1
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Fortea E, Lee S, Chadda R, Argyros Y, Sandal P, Mahoney-Kruszka R, Ciftci HD, Falzone ME, Huysmans G, Robertson JL, Boudker O, Accardi A. Structural basis of pH-dependent activation in a CLC transporter. Nat Struct Mol Biol 2024; 31:644-656. [PMID: 38279055 DOI: 10.1038/s41594-023-01210-5] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/16/2023] [Accepted: 12/22/2023] [Indexed: 01/28/2024]
Abstract
CLCs are dimeric chloride channels and anion/proton exchangers that regulate processes such as muscle contraction and endo-lysosome acidification. Common gating controls their activity; its closure simultaneously silences both protomers, and its opening allows them to independently transport ions. Mutations affecting common gating in human CLCs cause dominant genetic disorders. The structural rearrangements underlying common gating are unknown. Here, using single-particle cryo-electron microscopy, we show that the prototypical Escherichia coli CLC-ec1 undergoes large-scale rearrangements in activating conditions. The slow, pH-dependent remodeling of the dimer interface leads to the concerted opening of the intracellular H+ pathways and is required for transport. The more frequent formation of short water wires in the open H+ pathway enables Cl- pore openings. Mutations at disease-causing sites favor CLC-ec1 activation and accelerate common gate opening in the human CLC-7 exchanger. We suggest that the pH activation mechanism of CLC-ec1 is related to the common gating of CLC-7.
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Affiliation(s)
- Eva Fortea
- Department of Physiology and Biophysics, Weill Cornell Medical School, New York, NY, USA
- Department of Anesthesiology, Weill Cornell Medical School, New York, NY, USA
| | - Sangyun Lee
- Department of Anesthesiology, Weill Cornell Medical School, New York, NY, USA
| | - Rahul Chadda
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
| | - Yiorgos Argyros
- Department of Anesthesiology, Weill Cornell Medical School, New York, NY, USA
- Department of Biochemistry, Weill Cornell Medical School, New York, NY, USA
| | - Priyanka Sandal
- Department of Molecular Physiology and Biophysics, The University of Iowa, Iowa City, IA, USA
| | - Robyn Mahoney-Kruszka
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
| | - Hatice Didar Ciftci
- Department of Physiology and Biophysics, Weill Cornell Medical School, New York, NY, USA
- Tri-Institutional Training Program in Chemical Biology, New York, NY, USA
| | - Maria E Falzone
- Department of Anesthesiology, Weill Cornell Medical School, New York, NY, USA
- Department of Biochemistry, Weill Cornell Medical School, New York, NY, USA
| | - Gerard Huysmans
- Department of Physiology and Biophysics, Weill Cornell Medical School, New York, NY, USA
- Erasmus University, Jette, Belgium
| | - Janice L Robertson
- Department of Biochemistry and Molecular Biophysics, Washington University School of Medicine, St. Louis, MO, USA
| | - Olga Boudker
- Department of Physiology and Biophysics, Weill Cornell Medical School, New York, NY, USA.
- Howard Hughes Medical Institute, Chevy Chase, MD, USA.
| | - Alessio Accardi
- Department of Physiology and Biophysics, Weill Cornell Medical School, New York, NY, USA.
- Department of Anesthesiology, Weill Cornell Medical School, New York, NY, USA.
- Department of Biochemistry, Weill Cornell Medical School, New York, NY, USA.
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2
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Shi L, Yang C, Zhang M, Li K, Wang K, Jiao L, Liu R, Wang Y, Li M, Wang Y, Ma L, Hu S, Bian X. Dissecting the mechanism of atlastin-mediated homotypic membrane fusion at the single-molecule level. Nat Commun 2024; 15:2488. [PMID: 38509071 PMCID: PMC10954664 DOI: 10.1038/s41467-024-46919-z] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/27/2023] [Accepted: 03/13/2024] [Indexed: 03/22/2024] Open
Abstract
Homotypic membrane fusion of the endoplasmic reticulum (ER) is mediated by dynamin-like GTPase atlastin (ATL). This fundamental process relies on GTP-dependent domain rearrangements in the N-terminal region of ATL (ATLcyto), including the GTPase domain and three-helix bundle (3HB). However, its conformational dynamics during the GTPase cycle remain elusive. Here, we combine single-molecule FRET imaging and molecular dynamics simulations to address this conundrum. Different from the prevailing model, ATLcyto can form a loose crossover dimer upon GTP binding, which is tightened by GTP hydrolysis for membrane fusion. Furthermore, the α-helical motif between the 3HB and transmembrane domain, which is embedded in the surface of the lipid bilayer and self-associates in the crossover dimer, is required for ATL function. To recycle the proteins, Pi release, which disassembles the dimer, activates frequent relative movements between the GTPase domain and 3HB, and subsequent GDP dissociation alters the conformational preference of the ATLcyto monomer for entering the next reaction cycle. Finally, we found that two disease-causing mutations affect human ATL1 activity by destabilizing GTP binding-induced loose crossover dimer formation and the membrane-embedded helix, respectively. These results provide insights into ATL-mediated homotypic membrane fusion and the pathological mechanisms of related disease.
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Affiliation(s)
- Lijun Shi
- State Key Laboratory of Medicinal Chemical Biology, College of Life Sciences, Frontiers Science Center for Cell Responses, Nankai University, Tianjin, 300071, China
| | - Chenguang Yang
- National Laboratory for Condensed Matter Physics, Institute of Physics, Chinese Academy of Sciences, Beijing, 100190, China
- University of Chinese Academy of Sciences, Beijing, 100049, China
| | - Mingyuan Zhang
- College of Life Sciences, Zhejiang University, Hangzhou, 310027, China
| | - Kangning Li
- State Key Laboratory of Medicinal Chemical Biology, College of Life Sciences, Frontiers Science Center for Cell Responses, Nankai University, Tianjin, 300071, China
| | - Keying Wang
- College of Life Sciences, Zhejiang University, Hangzhou, 310027, China
| | - Li Jiao
- College of Life Sciences, Nankai University, Tianjin, 300071, China
| | - Ruming Liu
- College of Life Sciences, Nankai University, Tianjin, 300071, China
| | - Yunyun Wang
- State Key Laboratory of Medicinal Chemical Biology, College of Life Sciences, Frontiers Science Center for Cell Responses, Nankai University, Tianjin, 300071, China
| | - Ming Li
- National Laboratory for Condensed Matter Physics, Institute of Physics, Chinese Academy of Sciences, Beijing, 100190, China
| | - Yong Wang
- College of Life Sciences, Zhejiang University, Hangzhou, 310027, China.
- The Provincial International Science and Technology Cooperation Base on Engineering Biology, International Campus of Zhejiang University, Haining, 314400, China.
| | - Lu Ma
- National Laboratory for Condensed Matter Physics, Institute of Physics, Chinese Academy of Sciences, Beijing, 100190, China.
| | - Shuxin Hu
- National Laboratory for Condensed Matter Physics, Institute of Physics, Chinese Academy of Sciences, Beijing, 100190, China.
| | - Xin Bian
- State Key Laboratory of Medicinal Chemical Biology, College of Life Sciences, Frontiers Science Center for Cell Responses, Nankai University, Tianjin, 300071, China.
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3
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Sethi S, Wijesinghe KM, Dhakal S. Single-Molecule FRET-Based Multiplexed Detection. Methods Mol Biol 2024; 2744:183-195. [PMID: 38683319 DOI: 10.1007/978-1-0716-3581-0_11] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 05/01/2024]
Abstract
Single-molecule multiplexed detection is a high-promise toolkit for the expanding field of biosensing and molecular diagnostics. Among many single-molecule techniques available today for biomarker sensing including fluorescence, force, electrochemical, spectroscopic, barcoding, and other techniques, fluorescence-based approaches are arguably the most widely used methods due to their high sensitivity, selectivity, and readily available fluorophore-labeling schemes for a wide variety of biomolecules. However, multiplexed imaging using fluorescence techniques has proven to be challenging due to the sophisticated labeling schemes often requiring multiple FRET (fluorescence resonance energy transfer) pairs and/or excitation sources, which lead to overlapping signals and complicate data analysis. Here, we describe a single-molecule FRET method that enables multiplexed analysis while still using only one FRET pair, and thus the described approach is a significant step forward from conventional FRET methods.
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Affiliation(s)
- Srishty Sethi
- Department of Chemistry, Virginia Commonwealth University, Richmond, VA, USA
| | - Kalani M Wijesinghe
- Department of Chemistry, Virginia Commonwealth University, Richmond, VA, USA
| | - Soma Dhakal
- Department of Chemistry, Virginia Commonwealth University, Richmond, VA, USA.
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4
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Filius M, van Wee R, Joo C. Single-Molecule FRET X. Methods Mol Biol 2024; 2694:203-213. [PMID: 37824006 DOI: 10.1007/978-1-0716-3377-9_10] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 10/13/2023]
Abstract
Fluorescence resonance energy transfer (FRET) is a photophysical phenomenon that has been repurposed as a biophysical tool to measure nanometer distances. With FRET by DNA eXchange, or FRET X, many points of interest (POIs) in a single object can be probed, overcoming a major limitation of conventional single-molecule FRET. In FRET X, short fluorescently labeled DNA imager strands specifically and transiently bind their complementary docking strands on a target molecule, such that at most a single FRET pair is formed at each point in time and multiple POIs on a single molecule can be readily probed. Here, we describe the sample preparation, image acquisition, and data analysis for structural analysis of DNA nanostructures with FRET X.
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Affiliation(s)
- Mike Filius
- Department of BioNanoScience, Kavli Institute of Nanoscience, Delft University of Technology, Delft, The Netherlands
| | - Raman van Wee
- Department of BioNanoScience, Kavli Institute of Nanoscience, Delft University of Technology, Delft, The Netherlands
| | - Chirlmin Joo
- Department of BioNanoScience, Kavli Institute of Nanoscience, Delft University of Technology, Delft, The Netherlands.
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5
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Golfier S, Quail T, Brugués J. Single-Molecule Approaches to Study DNA Condensation. Methods Mol Biol 2024; 2740:1-19. [PMID: 38393466 DOI: 10.1007/978-1-0716-3557-5_1] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/25/2024]
Abstract
Proteins drive genome compartmentalization across different length scales. While the identities of these proteins have been well-studied, the physical mechanisms that drive genome organization have remained largely elusive. Studying these mechanisms is challenging owing to a lack of methodologies to parametrize physical models in cellular contexts. Furthermore, because of the complex, entangled, and dense nature of chromatin, conventional live imaging approaches often lack the spatial resolution to dissect these principles. In this chapter, we will describe how to image the interactions of λ-DNA with proteins under purified and cytoplasmic conditions. First, we will outline how to prepare biotinylated DNA, functionalize coverslips with biotin-conjugated poly-ethylene glycol (PEG), and assemble DNA microchannels compatible for the imaging of protein-DNA interactions using total internal fluorescence microscopy. Then we will describe experimental methods to image protein-DNA interactions in vitro and DNA loop extrusion using Xenopus laevis egg extracts.
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Affiliation(s)
- Stefan Golfier
- Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany
- Max Planck Institute for the Physics of Complex Systems, Dresden, Germany
- Center for Systems Biology Dresden, Dresden, Germany
- B CUBE, Center for Molecular Bioengineering, TU Dresden, Dresden, Germany
| | - Thomas Quail
- Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany
- Max Planck Institute for the Physics of Complex Systems, Dresden, Germany
- Center for Systems Biology Dresden, Dresden, Germany
- EMBL Heidelberg, Cell Biology and Biophysics Unit, Heidelberg, Germany
| | - Jan Brugués
- Max Planck Institute of Molecular Cell Biology and Genetics, Dresden, Germany.
- Max Planck Institute for the Physics of Complex Systems, Dresden, Germany.
- Center for Systems Biology Dresden, Dresden, Germany.
- Physics of Life, TU Dresden, Dresden, Germany.
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6
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Djaja N, Myong S. Protocol for single-molecule pull-down of fluorescently tagged oligomers from cell lysates. STAR Protoc 2023; 4:102716. [PMID: 37967012 PMCID: PMC10684877 DOI: 10.1016/j.xpro.2023.102716] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/29/2023] [Revised: 08/29/2023] [Accepted: 10/27/2023] [Indexed: 11/17/2023] Open
Abstract
Mutations in intrinsically disordered proteins drive the irreversible formation of pathological aggregates, a hallmark of neurodegenerative diseases. Here, we present a protocol to pull down fluorescently tagged proteins to characterize their basal oligomeric states. We describe steps for transfection and cell lysis, single-molecule slide preparation and pull-down, and oligomer dissolution. This protocol enables visualization of protein oligomers with single-molecule resolution. In addition, differences in oligomerization may provide insight on condensation or aggregation propensity in differing mutated or cell stress conditions. For complete details on the use and execution of this protocol, please refer to Djaja et al.1.
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Affiliation(s)
- Nathalie Djaja
- Program in Cell, Molecular, Developmental Biology, and Biophysics, Johns Hopkins University, Baltimore, MD 21218, USA; Department of Biology, Johns Hopkins University, Baltimore, MD 21218.
| | - Sua Myong
- Department of Biology, Johns Hopkins University, Baltimore, MD 21218; Department of Biophysics, Johns Hopkins University, Baltimore, MD 21218.
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7
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Oleinikov PD, Fedulova AS, Armeev GA, Motorin NA, Singh-Palchevskaia L, Sivkina AL, Feskin PG, Glukhov GS, Afonin DA, Komarova GA, Kirpichnikov MP, Studitsky VM, Feofanov AV, Shaytan AK. Interactions of Nucleosomes with Acidic Patch-Binding Peptides: A Combined Structural Bioinformatics, Molecular Modeling, Fluorescence Polarization, and Single-Molecule FRET Study. Int J Mol Sci 2023; 24:15194. [PMID: 37894874 PMCID: PMC10606924 DOI: 10.3390/ijms242015194] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/25/2023] [Revised: 10/08/2023] [Accepted: 10/11/2023] [Indexed: 10/29/2023] Open
Abstract
In eukaryotic organisms, genomic DNA associates with histone proteins to form nucleosomes. Nucleosomes provide a basis for genome compaction, epigenetic markup, and mediate interactions of nuclear proteins with their target DNA loci. A negatively charged (acidic) patch located on the H2A-H2B histone dimer is a characteristic feature of the nucleosomal surface. The acidic patch is a common site in the attachment of various chromatin proteins, including viral ones. Acidic patch-binding peptides present perspective compounds that can be used to modulate chromatin functioning by disrupting interactions of nucleosomes with natural proteins or alternatively targeting artificial moieties to the nucleosomes, which may be beneficial for the development of new therapeutics. In this work, we used several computational and experimental techniques to improve our understanding of how peptides may bind to the acidic patch and what are the consequences of their binding. Through extensive analysis of the PDB database, histone sequence analysis, and molecular dynamic simulations, we elucidated common binding patterns and key interactions that stabilize peptide-nucleosome complexes. Through MD simulations and FRET measurements, we characterized changes in nucleosome dynamics conferred by peptide binding. Using fluorescence polarization and gel electrophoresis, we evaluated the affinity and specificity of the LANA1-22 peptide to DNA and nucleosomes. Taken together, our study provides new insights into the different patterns of intermolecular interactions that can be employed by natural and designed peptides to bind to nucleosomes, and the effects of peptide binding on nucleosome dynamics and stability.
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Affiliation(s)
- Pavel D. Oleinikov
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
| | | | - Grigoriy A. Armeev
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
| | - Nikita A. Motorin
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
| | | | - Anastasiia L. Sivkina
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
- Laboratory of Structural-Functional Organization of Chromosomes, Institute of Gene Biology, Russian Academy of Sciences, 119334 Moscow, Russia
| | - Pavel G. Feskin
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
| | - Grigory S. Glukhov
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
- Faculty of Biology, MSU-BIT Shenzhen University, Shenzhen 518172, China
| | - Dmitry A. Afonin
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
| | - Galina A. Komarova
- Department of Physics, Lomonosov Moscow State University, 119234 Moscow, Russia
| | - Mikhail P. Kirpichnikov
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
- Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, 117997 Moscow, Russia
| | - Vasily M. Studitsky
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
- Fox Chase Cancer Center, Philadelphia, PA 19111, USA
| | - Alexey V. Feofanov
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
| | - Alexey K. Shaytan
- Department of Biology, Lomonosov Moscow State University, 119234 Moscow, Russia
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8
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Kim YH, Kim N, Okafor I, Choi S, Min S, Lee J, Bae SM, Choi K, Choi J, Harihar V, Kim Y, Kim JS, Kleinstiver BP, Lee JK, Ha T, Kim HH. Sniper2L is a high-fidelity Cas9 variant with high activity. Nat Chem Biol 2023:10.1038/s41589-023-01279-5. [PMID: 36894722 PMCID: PMC10374439 DOI: 10.1038/s41589-023-01279-5] [Citation(s) in RCA: 6] [Impact Index Per Article: 6.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/11/2022] [Accepted: 02/02/2023] [Indexed: 03/11/2023]
Abstract
Although several high-fidelity SpCas9 variants have been reported, it has been observed that this increased specificity is associated with reduced on-target activity, limiting the applications of the high-fidelity variants when efficient genome editing is required. Here, we developed an improved version of Sniper-Cas9, Sniper2L, which represents an exception to this trade-off trend as it showed higher specificity with retained high activity. We evaluated Sniper2L activities at a large number of target sequences and developed DeepSniper, a deep learning model that can predict the activity of Sniper2L. We also confirmed that Sniper2L can induce highly efficient and specific editing at a large number of target sequences when it is delivered as a ribonucleoprotein complex. Mechanically, the high specificity of Sniper2L originates from its superior ability to avoid unwinding a target DNA containing even a single mismatch. We envision that Sniper2L will be useful when efficient and specific genome editing is required.
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Affiliation(s)
- Young-Hoon Kim
- Toolgen, Seoul, Republic of Korea.,Department of Pharmacology, Yonsei University College of Medicine, Seoul, Republic of Korea.,Graduate Program of Biomedical Engineering, Yonsei University College of Medicine, Seoul, Republic of Korea.,Graduate Program of NanoScience and Technology, Yonsei University, Seoul, Republic of Korea
| | - Nahye Kim
- Department of Pharmacology, Yonsei University College of Medicine, Seoul, Republic of Korea.,Graduate School of Medical Science, Brain Korea 21 Project, Yonsei University College of Medicine, Seoul, Republic of Korea
| | - Ikenna Okafor
- Department of Biology, Johns Hopkins University, Baltimore, MD, USA
| | - Sungchul Choi
- Department of Pharmacology, Yonsei University College of Medicine, Seoul, Republic of Korea
| | | | | | | | | | - Janice Choi
- Department of Biophysics, Johns Hopkins University, Baltimore, MD, USA
| | - Vinayak Harihar
- Department of Biophysics, Johns Hopkins University, Baltimore, MD, USA
| | | | - Jin-Soo Kim
- Department of Biochemistry and NUS Synthetic Biology for Clinical & Technological Innovation (SynCTI), National University of Singapore, Singapore, Singapore
| | - Benjamin P Kleinstiver
- Center for Genomic Medicine, Massachusetts General Hospital, Boston, MA, USA.,Department of Pathology, Massachusetts General Hospital, Boston, MA, USA.,Department of Pathology, Harvard Medical School, Boston, MA, USA
| | | | - Taekjip Ha
- Department of Biophysics, Johns Hopkins University, Baltimore, MD, USA. .,Department of Biophysics and Biophysical Chemistry, Johns Hopkins University, Baltimore, MD, USA. .,Department of Biomedical Engineering, Johns Hopkins University, Baltimore, MD, USA. .,Howard Hughes Medical Institute, Baltimore, MD, USA.
| | - Hyongbum Henry Kim
- Department of Pharmacology, Yonsei University College of Medicine, Seoul, Republic of Korea. .,Graduate Program of NanoScience and Technology, Yonsei University, Seoul, Republic of Korea. .,Graduate School of Medical Science, Brain Korea 21 Project, Yonsei University College of Medicine, Seoul, Republic of Korea. .,Center for Nanomedicine, Institute for Basic Science, Seoul, Republic of Korea. .,Yonsei-Institute for Basic Science Institute, Yonsei University, Seoul, Republic of Korea. .,Severance Biomedical Science Institute, Yonsei University College of Medicine, Seoul, Republic of Korea. .,Institute for Immunology and Immunological Diseases, Yonsei University College of Medicine, Seoul, Republic of Korea.
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9
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Matamoros M, Ng XW, Brettmann JB, Piston DW, Nichols CG. Conformational plasticity of NaK2K and TREK2 potassium channel selectivity filters. Nat Commun 2023; 14:89. [PMID: 36609575 PMCID: PMC9822992 DOI: 10.1038/s41467-022-35756-7] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 05/18/2022] [Accepted: 12/23/2022] [Indexed: 01/07/2023] Open
Abstract
The K+ channel selectivity filter (SF) is defined by TxGYG amino acid sequences that generate four identical K+ binding sites (S1-S4). Only two sites (S3, S4) are present in the non-selective bacterial NaK channel, but a four-site K+-selective SF is obtained by mutating the wild-type TVGDGN SF sequence to a canonical K+ channel TVGYGD sequence (NaK2K mutant). Using single molecule FRET (smFRET), we show that the SF of NaK2K, but not of non-selective NaK, is ion-dependent, with the constricted SF configuration stabilized in high K+ conditions. Patch-clamp electrophysiology and non-canonical fluorescent amino acid incorporation show that NaK2K selectivity is reduced by crosslinking to limit SF conformational movement. Finally, the eukaryotic K+ channel TREK2 SF exhibits essentially identical smFRET-reported ion-dependent conformations as in prokaryotic K+ channels. Our results establish the generality of K+-induced SF conformational stability across the K+ channel superfamily, and introduce an approach to study manipulation of channel selectivity.
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Affiliation(s)
- Marcos Matamoros
- Center for Investigation of Membrane Excitability Diseases, Washington University School of Medicine, St. Louis, MO, USA
- Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO, USA
| | - Xue Wen Ng
- Center for Investigation of Membrane Excitability Diseases, Washington University School of Medicine, St. Louis, MO, USA
- Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO, USA
| | - Joshua B Brettmann
- Center for Investigation of Membrane Excitability Diseases, Washington University School of Medicine, St. Louis, MO, USA
- Millipore-Sigma Inc., St. Louis, MO, USA
| | - David W Piston
- Center for Investigation of Membrane Excitability Diseases, Washington University School of Medicine, St. Louis, MO, USA
- Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO, USA
| | - Colin G Nichols
- Center for Investigation of Membrane Excitability Diseases, Washington University School of Medicine, St. Louis, MO, USA.
- Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO, USA.
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10
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Ganser LR, Ge Y, Myong S. Single-Molecule Fluorescence Methods to Study Protein-RNA Interactions Underlying Biomolecular Condensates. Methods Mol Biol 2023; 2563:149-160. [PMID: 36227472 DOI: 10.1007/978-1-0716-2663-4_7] [Citation(s) in RCA: 4] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/16/2023]
Abstract
Many biomolecular condensates, including nucleoli and stress granules, form via dynamic multivalent protein-protein and protein-RNA interactions. These molecular interactions nucleate liquid-liquid phase separation (LLPS) and determine condensate properties, such as size and fluidity. Here, we outline the experimental procedures for single-molecule fluorescence experiments to probe protein-RNA interactions underlying LLPS. The experiments include single-molecule Förster (Fluorescence) resonance energy transfer (smFRET) to monitor protein-induced conformational changes in the RNA, protein-induced fluorescence enhancement (PIFE) to measure protein-RNA encounters, and single-molecule nucleation experiments to quantify the association and buildup of proteins on a nucleating RNA. Together, these experiments provide complementary approaches to elucidate a molecular view of the protein-RNA interactions that drive ribonucleoprotein condensate formation.
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Affiliation(s)
- Laura R Ganser
- Department of Biophysics, Johns Hopkins University, Baltimore, MD, USA
| | - Yingda Ge
- Department of Biophysics, Johns Hopkins University, Baltimore, MD, USA
| | - Sua Myong
- Department of Biophysics, Johns Hopkins University, Baltimore, MD, USA.
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11
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Choosing the Probe for Single-Molecule Fluorescence Microscopy. Int J Mol Sci 2022; 23:ijms232314949. [PMID: 36499276 PMCID: PMC9735909 DOI: 10.3390/ijms232314949] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/26/2022] [Revised: 11/18/2022] [Accepted: 11/24/2022] [Indexed: 12/03/2022] Open
Abstract
Probe choice in single-molecule microscopy requires deeper evaluations than those adopted for less sensitive fluorescence microscopy studies. Indeed, fluorophore characteristics can alter or hide subtle phenomena observable at the single-molecule level, wasting the potential of the sophisticated instrumentation and algorithms developed for advanced single-molecule applications. There are different reasons for this, linked, e.g., to fluorophore aspecific interactions, brightness, photostability, blinking, and emission and excitation spectra. In particular, these spectra and the excitation source are interdependent, and the latter affects the autofluorescence of sample substrate, medium, and/or biological specimen. Here, we review these and other critical points for fluorophore selection in single-molecule microscopy. We also describe the possible kinds of fluorophores and the microscopy techniques based on single-molecule fluorescence. We explain the importance and impact of the various issues in fluorophore choice, and discuss how this can become more effective and decisive for increasingly demanding experiments in single- and multiple-color applications.
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12
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Malinina DK, Sivkina AL, Korovina AN, McCullough LL, Formosa T, Kirpichnikov MP, Studitsky VM, Feofanov AV. Hmo1 Protein Affects the Nucleosome Structure and Supports the Nucleosome Reorganization Activity of Yeast FACT. Cells 2022; 11:cells11192931. [PMID: 36230893 PMCID: PMC9564320 DOI: 10.3390/cells11192931] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/16/2022] [Revised: 09/06/2022] [Accepted: 09/15/2022] [Indexed: 12/05/2022] Open
Abstract
Yeast Hmo1 is a high mobility group B (HMGB) protein that participates in the transcription of ribosomal protein genes and rDNA, and also stimulates the activities of some ATP-dependent remodelers. Hmo1 binds both DNA and nucleosomes and has been proposed to be a functional yeast analog of mammalian linker histones. We used EMSA and single particle Förster resonance energy transfer (spFRET) microscopy to characterize the effects of Hmo1 on nucleosomes alone and with the histone chaperone FACT. Hmo1 induced a significant increase in the distance between the DNA gyres across the nucleosomal core, and also caused the separation of linker segments. This was opposite to the effect of the linker histone H1, which enhanced the proximity of linkers. Similar to Nhp6, another HMGB factor, Hmo1, was able to support large-scale, ATP-independent, reversible unfolding of nucleosomes by FACT in the spFRET assay and partially support FACT function in vivo. However, unlike Hmo1, Nhp6 alone does not affect nucleosome structure. These results suggest physiological roles for Hmo1 that are distinct from Nhp6 and possibly from other HMGB factors and linker histones, such as H1.
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Affiliation(s)
- Daria K. Malinina
- Biology Faculty, Lomonosov Moscow State University, 119992 Moscow, Russia
| | | | - Anna N. Korovina
- Biology Faculty, Lomonosov Moscow State University, 119992 Moscow, Russia
| | - Laura L. McCullough
- Department of Biochemistry, University of Utah School of Medicine, Salt Lake City, UT 84132, USA
| | - Tim Formosa
- Department of Biochemistry, University of Utah School of Medicine, Salt Lake City, UT 84132, USA
| | - Mikhail P. Kirpichnikov
- Biology Faculty, Lomonosov Moscow State University, 119992 Moscow, Russia
- Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, 117997 Moscow, Russia
| | - Vasily M. Studitsky
- Biology Faculty, Lomonosov Moscow State University, 119992 Moscow, Russia
- Fox Chase Cancer Center, Philadelphia, PA 19111, USA
- Correspondence: (V.M.S.); (A.V.F.)
| | - Alexey V. Feofanov
- Biology Faculty, Lomonosov Moscow State University, 119992 Moscow, Russia
- Shemyakin-Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences, 117997 Moscow, Russia
- Correspondence: (V.M.S.); (A.V.F.)
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13
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Carcamo CC, Poyton MF, Ranjan A, Park G, Louder RK, Feng XA, Kim JM, Dzu T, Wu C, Ha T. ATP binding facilitates target search of SWR1 chromatin remodeler by promoting one-dimensional diffusion on DNA. eLife 2022; 11:e77352. [PMID: 35876491 PMCID: PMC9365391 DOI: 10.7554/elife.77352] [Citation(s) in RCA: 9] [Impact Index Per Article: 4.5] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/25/2022] [Accepted: 07/22/2022] [Indexed: 12/01/2022] Open
Abstract
One-dimensional (1D) target search is a well-characterized phenomenon for many DNA-binding proteins but is poorly understood for chromatin remodelers. Herein, we characterize the 1D scanning properties of SWR1, a conserved yeast chromatin remodeler that performs histone exchange on +1 nucleosomes adjacent to a nucleosome-depleted region (NDR) at gene promoters. We demonstrate that SWR1 has a kinetic binding preference for DNA of NDR length as opposed to gene-body linker length DNA. Using single and dual color single-particle tracking on DNA stretched with optical tweezers, we directly observe SWR1 diffusion on DNA. We found that various factors impact SWR1 scanning, including ATP which promotes diffusion through nucleotide binding rather than ATP hydrolysis. A DNA-binding subunit, Swc2, plays an important role in the overall diffusive behavior of the complex, as the subunit in isolation retains similar, although faster, scanning properties as the whole remodeler. ATP-bound SWR1 slides until it encounters a protein roadblock, of which we tested dCas9 and nucleosomes. The median diffusion coefficient, 0.024 μm2/s, in the regime of helical sliding, would mediate rapid encounter of NDR-flanking nucleosomes at length scales found in cellular chromatin.
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Affiliation(s)
- Claudia C Carcamo
- Department of Biophysics and Biophysical Chemistry, Johns Hopkins UniversityBaltimoreUnited States
| | - Matthew F Poyton
- Department of Biophysics and Biophysical Chemistry, Johns Hopkins UniversityBaltimoreUnited States
| | - Anand Ranjan
- Department of Biology, Johns Hopkins UniversityBaltimoreUnited States
| | - Giho Park
- Department of Biology, Johns Hopkins UniversityBaltimoreUnited States
| | - Robert K Louder
- Department of Biology, Johns Hopkins UniversityBaltimoreUnited States
| | - Xinyu A Feng
- Department of Biophysics and Biophysical Chemistry, Johns Hopkins UniversityBaltimoreUnited States
| | - Jee Min Kim
- Department of Biology, Johns Hopkins UniversityBaltimoreUnited States
| | - Thuc Dzu
- Department of Biology, Johns Hopkins UniversityBaltimoreUnited States
| | - Carl Wu
- Department of Biology, Johns Hopkins UniversityBaltimoreUnited States
| | - Taekjip Ha
- Department of Biophysics and Biophysical Chemistry, Johns Hopkins UniversityBaltimoreUnited States
- Howard Hughes Medical InstituteBaltimoreUnited States
- Johns Hopkins University, Department of Biomedical EngineeringBaltimoreUnited States
- Johns Hopkins University, Department of BiophysicsBaltimoreUnited States
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14
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Rhine K, Myong S. Single molecule probing of disordered RNA binding proteins. STAR Protoc 2022; 3:101131. [PMID: 35128475 PMCID: PMC8808284 DOI: 10.1016/j.xpro.2022.101131] [Citation(s) in RCA: 0] [Impact Index Per Article: 0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 10/26/2022] Open
Abstract
Liquid-liquid phase separation of intrinsically disordered proteins is known to underlie diverse pathologies such as neurodegeneration, cancer, and aging. The nucleation step of condensate formation is of critical importance for understanding how healthy and disease-associated condensates differ. Here, we describe four orthogonal single-molecule techniques that enable molecular tracking of the RNA-protein interaction, RNA-induced oligomerization, and kinetics of nucleation. These approaches allow researchers to directly interrogate the initial steps of liquid-liquid phase separation. For complete details on the use and execution of this profile, please refer to Niaki et al. (2020), Rhine et al. (2020), and Rhine et al. (2022).
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Affiliation(s)
- Kevin Rhine
- Program in Cell, Molecular, Developmental Biology and Biophysics, Johns Hopkins University, Baltimore, MD 21218, USA.,Department of Biology, Johns Hopkins University, Baltimore, MD 21218, USA
| | - Sua Myong
- Program in Cell, Molecular, Developmental Biology and Biophysics, Johns Hopkins University, Baltimore, MD 21218, USA.,Department of Biophysics, Johns Hopkins University, Baltimore, MD 21218, USA.,Physics Frontier Center (Center for the Physics of Living Cells), University of Illinois Urbana-Champaign, Urbana, IL 61801, USA
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15
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Paul T, Myong S. Protocol for generation and regeneration of PEG-passivated slides for single-molecule measurements. STAR Protoc 2022; 3:101152. [PMID: 35146451 PMCID: PMC8819390 DOI: 10.1016/j.xpro.2022.101152] [Citation(s) in RCA: 8] [Impact Index Per Article: 4.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/21/2022] Open
Abstract
Single-molecule fluorescence detection by total internal reflection microscope requires surface passivation by polyethylene glycol (PEG) coating, which is labor intensive and is only good for one or two experiments. Here, we present an efficient and reliable protocol for generating and regenerating the PEG surface for multiple rounds of experiments (∼5–10 times) in the same channel. This protocol is very simple, robust, rapid, and versatile; i.e., multiple strategies can be implemented to regenerate different layers of surface. The regeneration strategy saves time, improves the cost effectiveness, and enhances the efficiency of single-molecule experiments. For complete details on the use and execution of this profile, please refer to Paul et al. (2021a). Regeneration of PEG-passivated slide is simple, quick, and cost effective Multiple experiments can be performed in a single channel Different strategies are implemented for different level of regeneration Regeneration leads to highly reproducible results
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Affiliation(s)
- Tapas Paul
- Department of Biophysics, Johns Hopkins University, Baltimore, MD 21218, USA
| | - Sua Myong
- Department of Biophysics, Johns Hopkins University, Baltimore, MD 21218, USA.,Physics Frontier Center, Center for the Physics of Living Cells, University of Illinois Urbana-Champaign, Urbana, IL 61801, USA
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16
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Nasser M, Meller A. Lifetime-based analysis of binary fluorophores mixtures in the low photon count limit. iScience 2022; 25:103554. [PMID: 34977508 PMCID: PMC8689154 DOI: 10.1016/j.isci.2021.103554] [Citation(s) in RCA: 2] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 07/18/2021] [Revised: 10/18/2021] [Accepted: 11/30/2021] [Indexed: 11/28/2022] Open
Abstract
Single biomolecule sensing often requires the quantification of multiple fluorescent species. Here, we theoretically and experimentally use time-resolved fluorescence via Time Correlated Single Photon Counting (TCSPC) to accurately quantify fluorescent species with similar chromatic signatures. A modified maximum likelihood estimator is introduced to include two fluorophore species, with compensation of the instrument response function. We apply this algorithm to simulated data of a simplified two-fluorescent species model, as well as to experimental data of fluorophores' mixtures and to a model protein, doubly labeled with different fluorophores' ratio. We show that 100 to 200 photons per fluorophore, in a 10-ms timescale, are sufficient to provide an accurate estimation of the dyes' ratio on the model protein. Our results provide estimation for the desired photon integration time toward implementation of TCSPC in systems with fast occurring events, such as translocation of biomolecules through nanopores or single-molecule burst analyses experiments. Exact ratios of emission-similar dyes in binary mixtures were quantified by TCSPC MLE-based analysis with IRF compensation was implemented for two fluorescent dyes Dual dye bioconjugation on a model protein was quantified at limited photon counts
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Affiliation(s)
- Maisa Nasser
- Department of Biomedical Engineering, Technion - IIT, Haifa 32000, Israel
| | - Amit Meller
- Department of Biomedical Engineering, Technion - IIT, Haifa 32000, Israel
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17
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Fairlamb MS, Whitaker AM, Bain FE, Spies M, Freudenthal BD. Construction of a Three-Color Prism-Based TIRF Microscope to Study the Interactions and Dynamics of Macromolecules. BIOLOGY 2021; 10:biology10070571. [PMID: 34201434 PMCID: PMC8301196 DOI: 10.3390/biology10070571] [Citation(s) in RCA: 3] [Impact Index Per Article: 1.0] [Reference Citation Analysis] [Abstract] [Key Words] [Grants] [Track Full Text] [Download PDF] [Figures] [Subscribe] [Scholar Register] [Received: 05/11/2021] [Revised: 06/08/2021] [Accepted: 06/15/2021] [Indexed: 02/05/2023]
Abstract
Simple Summary Prism-based single-molecule total internal reflection fluorescence (prismTIRF) microscopes are excellent tools for studying macromolecular dynamics and interactions. Here, we provide an easy-to-follow guide for the design, assembly, and operation of a three-color prismTIRF microscope using commercially available components with the hope of assisting those who aim to implement TIRF imaging techniques in their laboratory. Abstract Single-molecule total internal reflection fluorescence (TIRF) microscopy allows for the real-time visualization of macromolecular dynamics and complex assembly. Prism-based TIRF microscopes (prismTIRF) are relatively simple to operate and can be easily modulated to fit the needs of a wide variety of experimental applications. While building a prismTIRF microscope without expert assistance can pose a significant challenge, the components needed to build a prismTIRF microscope are relatively affordable and, with some guidance, the assembly can be completed by a determined novice. Here, we provide an easy-to-follow guide for the design, assembly, and operation of a three-color prismTIRF microscope which can be utilized for the study of macromolecular complexes, including the multi-component protein–DNA complexes responsible for DNA repair, replication, and transcription. Our hope is that this article can assist laboratories that aspire to implement single-molecule TIRF techniques, and consequently expand the application of this technology.
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Affiliation(s)
- Max S. Fairlamb
- Department of Biochemistry and Molecular Biology and Department of Cancer Biology, University of Kansas Medical Center, Kansas City, KS 66160, USA; (M.S.F.); (A.M.W.)
| | - Amy M. Whitaker
- Department of Biochemistry and Molecular Biology and Department of Cancer Biology, University of Kansas Medical Center, Kansas City, KS 66160, USA; (M.S.F.); (A.M.W.)
| | - Fletcher E. Bain
- Department of Biochemistry and Molecular Biology, University of Iowa Carver College of Medicine, 51 Newton Road, Iowa City, IA 52242, USA; (F.E.B.); (M.S.)
| | - Maria Spies
- Department of Biochemistry and Molecular Biology, University of Iowa Carver College of Medicine, 51 Newton Road, Iowa City, IA 52242, USA; (F.E.B.); (M.S.)
| | - Bret D. Freudenthal
- Department of Biochemistry and Molecular Biology and Department of Cancer Biology, University of Kansas Medical Center, Kansas City, KS 66160, USA; (M.S.F.); (A.M.W.)
- Correspondence:
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18
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Lerner E, Barth A, Hendrix J, Ambrose B, Birkedal V, Blanchard SC, Börner R, Sung Chung H, Cordes T, Craggs TD, Deniz AA, Diao J, Fei J, Gonzalez RL, Gopich IV, Ha T, Hanke CA, Haran G, Hatzakis NS, Hohng S, Hong SC, Hugel T, Ingargiola A, Joo C, Kapanidis AN, Kim HD, Laurence T, Lee NK, Lee TH, Lemke EA, Margeat E, Michaelis J, Michalet X, Myong S, Nettels D, Peulen TO, Ploetz E, Razvag Y, Robb NC, Schuler B, Soleimaninejad H, Tang C, Vafabakhsh R, Lamb DC, Seidel CAM, Weiss S. FRET-based dynamic structural biology: Challenges, perspectives and an appeal for open-science practices. eLife 2021; 10:e60416. [PMID: 33779550 PMCID: PMC8007216 DOI: 10.7554/elife.60416] [Citation(s) in RCA: 125] [Impact Index Per Article: 41.7] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/29/2020] [Accepted: 02/09/2021] [Indexed: 12/18/2022] Open
Abstract
Single-molecule FRET (smFRET) has become a mainstream technique for studying biomolecular structural dynamics. The rapid and wide adoption of smFRET experiments by an ever-increasing number of groups has generated significant progress in sample preparation, measurement procedures, data analysis, algorithms and documentation. Several labs that employ smFRET approaches have joined forces to inform the smFRET community about streamlining how to perform experiments and analyze results for obtaining quantitative information on biomolecular structure and dynamics. The recent efforts include blind tests to assess the accuracy and the precision of smFRET experiments among different labs using various procedures. These multi-lab studies have led to the development of smFRET procedures and documentation, which are important when submitting entries into the archiving system for integrative structure models, PDB-Dev. This position paper describes the current 'state of the art' from different perspectives, points to unresolved methodological issues for quantitative structural studies, provides a set of 'soft recommendations' about which an emerging consensus exists, and lists openly available resources for newcomers and seasoned practitioners. To make further progress, we strongly encourage 'open science' practices.
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Affiliation(s)
- Eitan Lerner
- Department of Biological Chemistry, The Alexander Silberman Institute of Life Sciences, and The Center for Nanoscience and Nanotechnology, Faculty of Mathematics & Science, The Edmond J. Safra Campus, The Hebrew University of JerusalemJerusalemIsrael
| | - Anders Barth
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-UniversitätDüsseldorfGermany
| | - Jelle Hendrix
- Dynamic Bioimaging Lab, Advanced Optical Microscopy Centre and Biomedical Research Institute (BIOMED), Hasselt UniversityDiepenbeekBelgium
| | - Benjamin Ambrose
- Department of Chemistry, University of SheffieldSheffieldUnited Kingdom
| | - Victoria Birkedal
- Department of Chemistry and iNANO center, Aarhus UniversityAarhusDenmark
| | - Scott C Blanchard
- Department of Structural Biology, St. Jude Children's Research HospitalMemphisUnited States
| | - Richard Börner
- Laserinstitut HS Mittweida, University of Applied Science MittweidaMittweidaGermany
| | - Hoi Sung Chung
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of HealthBethesdaUnited States
| | - Thorben Cordes
- Physical and Synthetic Biology, Faculty of Biology, Ludwig-Maximilians-Universität MünchenPlanegg-MartinsriedGermany
| | - Timothy D Craggs
- Department of Chemistry, University of SheffieldSheffieldUnited Kingdom
| | - Ashok A Deniz
- Department of Integrative Structural and Computational Biology, The Scripps Research InstituteLa JollaUnited States
| | - Jiajie Diao
- Department of Cancer Biology, University of Cincinnati School of MedicineCincinnatiUnited States
| | - Jingyi Fei
- Department of Biochemistry and Molecular Biology and The Institute for Biophysical Dynamics, University of ChicagoChicagoUnited States
| | - Ruben L Gonzalez
- Department of Chemistry, Columbia UniversityNew YorkUnited States
| | - Irina V Gopich
- Laboratory of Chemical Physics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of HealthBethesdaUnited States
| | - Taekjip Ha
- Department of Biophysics and Biophysical Chemistry, Department of Biomedical Engineering, Johns Hopkins University School of Medicine, Howard Hughes Medical InstituteBaltimoreUnited States
| | - Christian A Hanke
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-UniversitätDüsseldorfGermany
| | - Gilad Haran
- Department of Chemical and Biological Physics, Weizmann Institute of ScienceRehovotIsrael
| | - Nikos S Hatzakis
- Department of Chemistry & Nanoscience Centre, University of CopenhagenCopenhagenDenmark
- Denmark Novo Nordisk Foundation Centre for Protein Research, Faculty of Health and Medical Sciences, University of CopenhagenCopenhagenDenmark
| | - Sungchul Hohng
- Department of Physics and Astronomy, and Institute of Applied Physics, Seoul National UniversitySeoulRepublic of Korea
| | - Seok-Cheol Hong
- Center for Molecular Spectroscopy and Dynamics, Institute for Basic Science and Department of Physics, Korea UniversitySeoulRepublic of Korea
| | - Thorsten Hugel
- Institute of Physical Chemistry and Signalling Research Centres BIOSS and CIBSS, University of FreiburgFreiburgGermany
| | - Antonino Ingargiola
- Department of Chemistry and Biochemistry, and Department of Physiology, University of California, Los AngelesLos AngelesUnited States
| | - Chirlmin Joo
- Department of BioNanoScience, Kavli Institute of Nanoscience, Delft University of TechnologyDelftNetherlands
| | - Achillefs N Kapanidis
- Biological Physics Research Group, Clarendon Laboratory, Department of Physics, University of OxfordOxfordUnited Kingdom
| | - Harold D Kim
- School of Physics, Georgia Institute of TechnologyAtlantaUnited States
| | - Ted Laurence
- Physical and Life Sciences Directorate, Lawrence Livermore National LaboratoryLivermoreUnited States
| | - Nam Ki Lee
- School of Chemistry, Seoul National UniversitySeoulRepublic of Korea
| | - Tae-Hee Lee
- Department of Chemistry, Pennsylvania State UniversityUniversity ParkUnited States
| | - Edward A Lemke
- Departments of Biology and Chemistry, Johannes Gutenberg UniversityMainzGermany
- Institute of Molecular Biology (IMB)MainzGermany
| | - Emmanuel Margeat
- Centre de Biologie Structurale (CBS), CNRS, INSERM, Universitié de MontpellierMontpellierFrance
| | | | - Xavier Michalet
- Department of Chemistry and Biochemistry, and Department of Physiology, University of California, Los AngelesLos AngelesUnited States
| | - Sua Myong
- Department of Biophysics, Johns Hopkins UniversityBaltimoreUnited States
| | - Daniel Nettels
- Department of Biochemistry and Department of Physics, University of ZurichZurichSwitzerland
| | - Thomas-Otavio Peulen
- Department of Bioengineering and Therapeutic Sciences, University of California, San FranciscoSan FranciscoUnited States
| | - Evelyn Ploetz
- Physical Chemistry, Department of Chemistry, Center for Nanoscience (CeNS), Center for Integrated Protein Science Munich (CIPSM) and Nanosystems Initiative Munich (NIM), Ludwig-Maximilians-UniversitätMünchenGermany
| | - Yair Razvag
- Department of Biological Chemistry, The Alexander Silberman Institute of Life Sciences, and The Center for Nanoscience and Nanotechnology, Faculty of Mathematics & Science, The Edmond J. Safra Campus, The Hebrew University of JerusalemJerusalemIsrael
| | - Nicole C Robb
- Warwick Medical School, University of WarwickCoventryUnited Kingdom
| | - Benjamin Schuler
- Department of Biochemistry and Department of Physics, University of ZurichZurichSwitzerland
| | - Hamid Soleimaninejad
- Biological Optical Microscopy Platform (BOMP), University of MelbourneParkvilleAustralia
| | - Chun Tang
- College of Chemistry and Molecular Engineering, PKU-Tsinghua Center for Life Sciences, Beijing National Laboratory for Molecular Sciences, Peking UniversityBeijingChina
| | - Reza Vafabakhsh
- Department of Molecular Biosciences, Northwestern UniversityEvanstonUnited States
| | - Don C Lamb
- Physical Chemistry, Department of Chemistry, Center for Nanoscience (CeNS), Center for Integrated Protein Science Munich (CIPSM) and Nanosystems Initiative Munich (NIM), Ludwig-Maximilians-UniversitätMünchenGermany
| | - Claus AM Seidel
- Lehrstuhl für Molekulare Physikalische Chemie, Heinrich-Heine-UniversitätDüsseldorfGermany
| | - Shimon Weiss
- Department of Chemistry and Biochemistry, and Department of Physiology, University of California, Los AngelesLos AngelesUnited States
- Department of Physiology, CaliforniaNanoSystems Institute, University of California, Los AngelesLos AngelesUnited States
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19
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Matamoros M, Nichols CG. Pore-forming transmembrane domains control ion selectivity and selectivity filter conformation in the KirBac1.1 potassium channel. J Gen Physiol 2021; 153:211923. [PMID: 33779689 PMCID: PMC8008366 DOI: 10.1085/jgp.202012683] [Citation(s) in RCA: 6] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 06/18/2020] [Revised: 11/23/2020] [Accepted: 03/09/2021] [Indexed: 01/10/2023] Open
Abstract
Potassium (K+) channels are membrane proteins with the remarkable ability to very selectively conduct K+ ions across the membrane. High-resolution structures have revealed that dehydrated K+ ions permeate through the narrowest region of the pore, formed by the backbone carbonyls of the signature selectivity filter (SF) sequence TxGYG. However, the existence of nonselective channels with similar SF sequences, as well as effects of mutations in other regions on selectivity, suggest that the SF is not the sole determinant of selectivity. We changed the selectivity of the KirBac1.1 channel by introducing mutations at residue I131 in transmembrane helix 2 (TM2). These mutations increase Na+ flux in the absence of K+ and introduce significant proton conductance. Consistent with K+ channel crystal structures, single-molecule FRET experiments show that the SF is conformationally constrained and stable in high-K+ conditions but undergoes transitions to dilated low-FRET states in high-Na+/low-K+ conditions. Relative to wild-type channels, I131M mutants exhibit marked shifts in the K+ and Na+ dependence of SF dynamics to higher K+ and lower Na+ concentrations. These results illuminate the role of I131, and potentially other structural elements outside the SF, in controlling ion selectivity, by suggesting that the physical interaction of these elements with the SF contributes to the relative stability of the constrained K+-induced SF configuration versus nonselective dilated conformations.
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Affiliation(s)
- Marcos Matamoros
- Center for Investigation of Membrane Excitability Diseases, and Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO
| | - Colin G Nichols
- Center for Investigation of Membrane Excitability Diseases, and Department of Cell Biology and Physiology, Washington University School of Medicine, St. Louis, MO
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20
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Kim SH, Kim H, Jeong H, Yoon TY. Encoding Multiple Virtual Signals in DNA Barcodes with Single-Molecule FRET. NANO LETTERS 2021; 21:1694-1701. [PMID: 33586985 DOI: 10.1021/acs.nanolett.0c04502] [Citation(s) in RCA: 4] [Impact Index Per Article: 1.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Track Full Text] [Subscribe] [Scholar Register] [Indexed: 05/13/2023]
Abstract
DNA barcoding provides a way to label a myriad of different biological molecules using the extreme programmability in DNA sequence synthesis. Fluorescence imaging is presumably the most easy-to-access method for DNA barcoding, yet large spectral overlaps between fluorescence dyes severely limit the numbers of barcodes that can be detected simultaneously. We here demonstrate the use of single-molecule fluorescence resonance energy transfer (FRET) to encode virtual signals in DNA barcodes using conventional two-color fluorescence microscopy. By optimizing imaging and biochemistry conditions for weak DNA hybridization events, we markedly enhanced accuracy in our determination of the single-molecule FRET efficiency exhibited by each binding event between DNA barcode sequences. This allowed us to unambiguously differentiate six DNA barcodes encoding different FRET values without involving any probe sequence exchanges. Our method can be directly incorporated with previous DNA barcode techniques, and may thus be widely adopted to expand the signal space of DNA barcoding.
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Affiliation(s)
- Sung Hyun Kim
- School of Biological Sciences and Institute for Molecular Biology and Genetics, Seoul National University, Seoul 08826, South Korea
| | - Hyunwoo Kim
- Department of Physics, Korea Advanced Institute of Science and Technology (KAIST), Daejeon 34141, South Korea
| | - Hawoong Jeong
- Department of Physics, Korea Advanced Institute of Science and Technology (KAIST), Daejeon 34141, South Korea
| | - Tae-Young Yoon
- School of Biological Sciences and Institute for Molecular Biology and Genetics, Seoul National University, Seoul 08826, South Korea
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21
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Ruijtenberg S, Sonneveld S, Cui TJ, Logister I, de Steenwinkel D, Xiao Y, MacRae IJ, Joo C, Tanenbaum ME. mRNA structural dynamics shape Argonaute-target interactions. Nat Struct Mol Biol 2020; 27:790-801. [PMID: 32661421 DOI: 10.1038/s41594-020-0461-1] [Citation(s) in RCA: 27] [Impact Index Per Article: 6.8] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/06/2020] [Accepted: 06/11/2020] [Indexed: 12/17/2022]
Abstract
Small interfering RNAs (siRNAs) promote RNA degradation in a variety of processes and have important clinical applications. siRNAs direct cleavage of target RNAs by guiding Argonaute2 (AGO2) to its target site. Target site accessibility is critical for AGO2-target interactions, but how target site accessibility is controlled in vivo is poorly understood. Here, we use live-cell single-molecule imaging in human cells to determine rate constants of the AGO2 cleavage cycle in vivo. We find that the rate-limiting step in mRNA cleavage frequently involves unmasking of target sites by translating ribosomes. Target site masking is caused by heterogeneous intramolecular RNA-RNA interactions, which can conceal target sites for many minutes in the absence of translation. Our results uncover how dynamic changes in mRNA structure shape AGO2-target recognition, provide estimates of mRNA folding and unfolding rates in vivo, and provide experimental evidence for the role of mRNA structural dynamics in control of mRNA-protein interactions.
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Affiliation(s)
- Suzan Ruijtenberg
- Oncode Institute, Hubrecht Institute-KNAW and University Medical Center Utrecht, Utrecht, the Netherlands.,Developmental Biology, Department of Biology, Faculty of Sciences, Utrecht University, Utrecht, the Netherlands
| | - Stijn Sonneveld
- Oncode Institute, Hubrecht Institute-KNAW and University Medical Center Utrecht, Utrecht, the Netherlands
| | - Tao Ju Cui
- Kavli Institute of NanoScience, Department of BioNanoScience, Delft University of Technology, Delft, the Netherlands
| | - Ive Logister
- Oncode Institute, Hubrecht Institute-KNAW and University Medical Center Utrecht, Utrecht, the Netherlands
| | - Dion de Steenwinkel
- Oncode Institute, Hubrecht Institute-KNAW and University Medical Center Utrecht, Utrecht, the Netherlands
| | - Yao Xiao
- Department of Integrative Structural and Computational Biology, The Scripps Research Institute, La Jolla, CA, USA
| | - Ian J MacRae
- Department of Integrative Structural and Computational Biology, The Scripps Research Institute, La Jolla, CA, USA
| | - Chirlmin Joo
- Kavli Institute of NanoScience, Department of BioNanoScience, Delft University of Technology, Delft, the Netherlands
| | - Marvin E Tanenbaum
- Oncode Institute, Hubrecht Institute-KNAW and University Medical Center Utrecht, Utrecht, the Netherlands.
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22
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Wang X, Gong J, Zhu L, Wang S, Yang X, Xu Y, Yang X, Ma C. Munc13 activates the Munc18-1/syntaxin-1 complex and enables Munc18-1 to prime SNARE assembly. EMBO J 2020; 39:e103631. [PMID: 32643828 DOI: 10.15252/embj.2019103631] [Citation(s) in RCA: 19] [Impact Index Per Article: 4.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/03/2019] [Revised: 05/20/2020] [Accepted: 06/02/2020] [Indexed: 11/09/2022] Open
Abstract
Priming of synaptic vesicles involves Munc13-catalyzed transition of the Munc18-1/syntaxin-1 complex to the SNARE complex in the presence of SNAP-25 and synaptobrevin-2; Munc13 drives opening of syntaxin-1 via the MUN domain while Munc18-1 primes SNARE assembly via domain 3a. However, the underlying mechanism remains unclear. In this study, we have identified a number of residues in domain 3a of Munc18-1 that are crucial for Munc13 and Munc18-1 actions in SNARE complex assembly and synaptic vesicle priming. Our results showed that two residues (Q301/K308) at the side of domain 3a mediate the interaction between the Munc18-1/syntaxin-1 complex and the MUN domain. This interaction enables the MUN domain to drive the opening of syntaxin-1 linker region, thereby leading to the extension of domain 3a and promoting synaptobrevin-2 binding. In addition, we identified two residues (K332/K333) at the bottom of domain 3a that mediate the interaction between Munc18-1 and the SNARE motif of syntaxin-1. This interaction ensures Munc18-1 to persistently associate with syntaxin-1 during the conformational change of syntaxin-1 from closed to open, which reinforces the role of Munc18-1 in templating SNARE assembly. Taken together, our data suggest a mechanism by which Munc13 activates the Munc18-1/syntaxin-1 complex and enables Munc18-1 to prime SNARE assembly.
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Affiliation(s)
- Xianping Wang
- Key Laboratory of Molecular Biophysics of the Ministry of Education, College of Life Science and Technology, Huazhong University of Science and Technology, Wuhan, China
| | - Jihong Gong
- Key Laboratory of Molecular Biophysics of the Ministry of Education, College of Life Science and Technology, Huazhong University of Science and Technology, Wuhan, China
| | - Le Zhu
- Key Laboratory of Molecular Biophysics of the Ministry of Education, College of Life Science and Technology, Huazhong University of Science and Technology, Wuhan, China
| | - Shen Wang
- Key Laboratory of Molecular Biophysics of the Ministry of Education, College of Life Science and Technology, Huazhong University of Science and Technology, Wuhan, China
| | - Xiaoyu Yang
- Key Laboratory of Molecular Biophysics of the Ministry of Education, College of Life Science and Technology, Huazhong University of Science and Technology, Wuhan, China
| | - Yuanyuan Xu
- Key Laboratory of Molecular Biophysics of the Ministry of Education, College of Life Science and Technology, Huazhong University of Science and Technology, Wuhan, China
| | - Xiaofei Yang
- Key Laboratory of Cognitive Science, Hubei Key Laboratory of Medical Information Analysis and Tumor Diagnosis & Treatment, Laboratory of Membrane Ion Channels and Medicine, College of Biomedical Engineering, South-Central University for Nationalities, Wuhan, China
| | - Cong Ma
- Key Laboratory of Molecular Biophysics of the Ministry of Education, College of Life Science and Technology, Huazhong University of Science and Technology, Wuhan, China.,Institute of Brain Research, Huazhong University of Science and Technology, Wuhan, China
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23
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R-loop induced G-quadruplex in non-template promotes transcription by successive R-loop formation. Nat Commun 2020; 11:3392. [PMID: 32636376 PMCID: PMC7341879 DOI: 10.1038/s41467-020-17176-7] [Citation(s) in RCA: 40] [Impact Index Per Article: 10.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 09/11/2019] [Accepted: 06/17/2020] [Indexed: 01/06/2023] Open
Abstract
G-quadruplex (G4) is a noncanonical secondary structure of DNA or RNA which can enhance or repress gene expression, yet the underlying molecular mechanism remains uncertain. Here we show that when positioned downstream of transcription start site, the orientation of potential G4 forming sequence (PQS), but not the sequence alters transcriptional output. Ensemble in vitro transcription assays indicate that PQS in the non-template increases mRNA production rate and yield. Using sequential single molecule detection stages, we demonstrate that while binding and initiation of T7 RNA polymerase is unchanged, the efficiency of elongation and the final mRNA output is higher when PQS is in the non-template. Strikingly, the enhanced elongation arises from the transcription-induced R-loop formation, which in turn generates G4 structure in the non-template. The G4 stabilized R-loop leads to increased transcription by a mechanism involving successive rounds of R-loop formation. G-quadruplex (G4) forming sequences are highly enriched in the human genome and function as important regulators of diverse range of biological processes. Here the authors show that while G4 structures on template strand block transcription, folding on the non-template strand enhances transcription by means of successive R-loop formation.
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24
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Wu WQ, Zhang ML, Song CP. A comprehensive evaluation of a typical plant telomeric G-quadruplex (G4) DNA reveals the dynamics of G4 formation, rearrangement, and unfolding. J Biol Chem 2020; 295:5461-5469. [PMID: 32184352 PMCID: PMC7170514 DOI: 10.1074/jbc.ra119.012383] [Citation(s) in RCA: 17] [Impact Index Per Article: 4.3] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/19/2019] [Revised: 03/16/2020] [Indexed: 11/06/2022] Open
Abstract
Telomeres are specific nucleoprotein structures that are located at the ends of linear eukaryotic chromosomes and play crucial roles in genomic stability. Telomere DNA consists of simple repeats of a short G-rich sequence: TTAGGG in mammals and TTTAGGG in most plants. In recent years, the mammalian telomeric G-rich repeats have been shown to form G-quadruplex (G4) structures, which are crucial for modulating telomere functions. Surprisingly, even though plant telomeres are essential for plant growth, development, and environmental adaptions, only few reports exist on plant telomeric G4 DNA (pTG4). Here, using bulk and single-molecule assays, including CD spectroscopy, and single-molecule FRET approaches, we comprehensively characterized the structure and dynamics of a typical plant telomeric sequence, d[GGG(TTTAGGG)3]. We found that this sequence can fold into mixed G4s in potassium, including parallel and antiparallel structures. We also directly detected intermediate dynamic transitions, including G-hairpin, parallel G-triplex, and antiparallel G-triplex structures. Moreover, we observed that pTG4 is unfolded by the AtRecQ2 helicase but not by AtRecQ3. The results of our work shed light on our understanding about the existence, topological structures, stability, intermediates, unwinding, and functions of pTG4.
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Affiliation(s)
- Wen-Qiang Wu
- State Key Laboratory of Crop Stress Adaptation and Improvement, Key Laboratory of Plant Stress Biology, School of Life Sciences, Henan University, Kaifeng 475001, China
| | - Ming-Li Zhang
- State Key Laboratory of Crop Stress Adaptation and Improvement, Key Laboratory of Plant Stress Biology, School of Life Sciences, Henan University, Kaifeng 475001, China
| | - Chun-Peng Song
- State Key Laboratory of Crop Stress Adaptation and Improvement, Key Laboratory of Plant Stress Biology, School of Life Sciences, Henan University, Kaifeng 475001, China.
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25
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Wang Y, Horáček M, Zijlstra P. Strong Plasmon Enhancement of the Saturation Photon Count Rate of Single Molecules. J Phys Chem Lett 2020; 11:1962-1969. [PMID: 32073865 PMCID: PMC7061331 DOI: 10.1021/acs.jpclett.0c00155] [Citation(s) in RCA: 6] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [Track Full Text] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/15/2020] [Accepted: 02/19/2020] [Indexed: 06/10/2023]
Abstract
Plasmon resonances have appeared as a promising method to boost the fluorescence intensity of single emitters. However, because research has focused on the enhancement at low excitation intensity, little is known about plasmon-fluorophore coupling near the point where the dye saturates. Here we study plasmon-enhanced fluorescence at a broad range of excitation intensities up to saturation. We adopt a novel DNA-mediated approach wherein dynamic single-molecule binding provides a controlled particle-fluorophore spacing, and dynamic rebinding circumvents artifacts due to photobleaching. We find that near saturation the maximum photon count rate is enhanced by more than 2 orders of magnitude at the optimal particle-fluorophore spacing, even for a dye with a high intrinsic quantum yield. We compare our results to a numerical model taking into account dye saturation. These experiments provide design rules to maximize the photon output of single emitters, which will open the door to studying fast dynamics in real time using single-molecule fluorescence.
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Affiliation(s)
- Yuyang Wang
- Department
of Applied Physics, Eindhoven University
of Technology, Postbus 513, 5600 MB Eindhoven, The Netherlands
- Institute
for Complex Molecular Systems, Eindhoven
University of Technology, Postbus 513, 5600 MB Eindhoven, The Netherlands
| | - Matěj Horáček
- Department
of Applied Physics, Eindhoven University
of Technology, Postbus 513, 5600 MB Eindhoven, The Netherlands
- Institute
for Complex Molecular Systems, Eindhoven
University of Technology, Postbus 513, 5600 MB Eindhoven, The Netherlands
| | - Peter Zijlstra
- Department
of Applied Physics, Eindhoven University
of Technology, Postbus 513, 5600 MB Eindhoven, The Netherlands
- Institute
for Complex Molecular Systems, Eindhoven
University of Technology, Postbus 513, 5600 MB Eindhoven, The Netherlands
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26
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Measurements of the self-assembly kinetics of individual viral capsids around their RNA genome. Proc Natl Acad Sci U S A 2019; 116:22485-22490. [PMID: 31570619 PMCID: PMC6842639 DOI: 10.1073/pnas.1909223116] [Citation(s) in RCA: 50] [Impact Index Per Article: 10.0] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/24/2023] Open
Abstract
Self-assembly is a process in which functional nanoscale structures build themselves, driven by Brownian motion and interactions between components. The term was originally coined to describe the formation of a viral capsid, the protein shell that protects the genome of a virus. Despite decades of study, how capsids self-assemble has remained a mystery, because there were no methods to measure the assembly kinetics of individual capsids. We surmount this obstacle using a sensitive microscopy technique based on laser interferometry. The measurements show that a small nucleus of proteins must form on the viral RNA before the capsid assembles. These results might help researchers design strategies to stop the assembly of pathogenic viruses or to build synthetic nanostructures. Self-assembly is widely used by biological systems to build functional nanostructures, such as the protein capsids of RNA viruses. But because assembly is a collective phenomenon involving many weakly interacting subunits and a broad range of timescales, measurements of the assembly pathways have been elusive. We use interferometric scattering microscopy to measure the assembly kinetics of individual MS2 bacteriophage capsids around MS2 RNA. By recording how many coat proteins bind to each of many individual RNA strands, we find that assembly proceeds by nucleation followed by monotonic growth. Our measurements reveal the assembly pathways in quantitative detail and also show their failure modes. We use these results to critically examine models of the assembly process.
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27
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Cui TJ, Klein M, Hegge JW, Chandradoss SD, van der Oost J, Depken M, Joo C. Argonaute bypasses cellular obstacles without hindrance during target search. Nat Commun 2019; 10:4390. [PMID: 31558728 PMCID: PMC6763497 DOI: 10.1038/s41467-019-12415-y] [Citation(s) in RCA: 8] [Impact Index Per Article: 1.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 02/11/2019] [Accepted: 09/09/2019] [Indexed: 12/13/2022] Open
Abstract
Argonaute (Ago) proteins are key players in both gene regulation (eukaryotes) and host defense (prokaryotes). Acting on single-stranded nucleic-acid substrates, Ago relies on base pairing between a small nucleic-acid guide and its complementary target sequences for specificity. To efficiently scan nucleic-acid chains for targets, Ago diffuses laterally along the substrate and must bypass secondary structures as well as protein barriers. Using single-molecule FRET in conjunction with kinetic modelling, we reveal that target scanning is mediated through loose protein-nucleic acid interactions, allowing Ago to slide short distances over secondary structures, as well as to bypass protein barriers via intersegmental transfer. Our combined single-molecule experiment and kinetic modelling approach may serve as a platform to dissect search processes and study the effect of sequence on search kinetics for other nucleic acid-guided proteins.
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Affiliation(s)
- Tao Ju Cui
- Kavli Institute of Nanoscience and Department of Bionanoscience, Delft University of Technology, Delft, The Netherlands
| | - Misha Klein
- Kavli Institute of Nanoscience and Department of Bionanoscience, Delft University of Technology, Delft, The Netherlands
| | - Jorrit W Hegge
- Laboratory of Microbiology, Department of Agrotechnology and Food Sciences, Wageningen University, Wageningen, The Netherlands
| | - Stanley D Chandradoss
- Kavli Institute of Nanoscience and Department of Bionanoscience, Delft University of Technology, Delft, The Netherlands.,Oxford NanoImaging, Oxford, UK
| | - John van der Oost
- Laboratory of Microbiology, Department of Agrotechnology and Food Sciences, Wageningen University, Wageningen, The Netherlands
| | - Martin Depken
- Kavli Institute of Nanoscience and Department of Bionanoscience, Delft University of Technology, Delft, The Netherlands.
| | - Chirlmin Joo
- Kavli Institute of Nanoscience and Department of Bionanoscience, Delft University of Technology, Delft, The Netherlands.
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28
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The essential elements for the noncovalent association of two DNA ends during NHEJ synapsis. Nat Commun 2019; 10:3588. [PMID: 31399561 PMCID: PMC6688983 DOI: 10.1038/s41467-019-11507-z] [Citation(s) in RCA: 50] [Impact Index Per Article: 10.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/07/2018] [Accepted: 07/17/2019] [Indexed: 01/13/2023] Open
Abstract
One of the most central questions about the repair of a double-strand DNA break (DSB) concerns how the two free DNA ends are brought together - a step called synapsis. Using single-molecule FRET (smFRET), we show here that both Ku plus XRCC4:DNA ligase IV are necessary and sufficient to achieve a flexible synapsis of blunt DNA ends, whereas either alone is not. Addition of XLF causes a transition to a close synaptic state, and maximum efficiency of close synapsis is achieved within 20 min. The promotion of close synapsis by XLF indicates a role that is independent of a filament structure, with action focused at the very ends of each duplex. DNA-PKcs is not required for the formation of either the flexible or close synaptic states. This model explains in biochemical terms the evolutionarily central synaptic role of Ku, X4L4, and XLF in NHEJ for all eukaryotes.
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29
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Cai B, Yu L, Sharum SR, Zhang K, Diao J. Single-vesicle measurement of protein-induced membrane tethering. Colloids Surf B Biointerfaces 2019; 177:267-273. [PMID: 30769228 DOI: 10.1016/j.colsurfb.2019.02.004] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/10/2018] [Revised: 01/28/2019] [Accepted: 02/04/2019] [Indexed: 12/30/2022]
Abstract
Functions of the proteins involved in membrane tethering, a crucial step in membrane trafficking, remain elusive due to the lack of effective tools to investigate protein-lipid interaction. To address this challenge, we introduce a method to study protein-induced membrane tethering via in vitro reconstitution of lipid vesicles, including detailed steps from the preparation of the PEGylated slides to the imaging of single vesicles. Furthermore, we demonstrate the measurement of protein-vesicle interaction in tethered vesicle pairs using two representative proteins, the cytoplasmic domain of synaptotagmin-1 (C2AB) and α-synuclein. Results from Förster (fluorescence) resonance energy transfer (FRET) reveal that membrane tethering is distinguished from membrane fusion. Single-vesicle measurement also allows for assessment of dose-dependent effects of proteins and ions on membrane tethering. We envision that the continuous development of advanced techniques in the single-vesicle measurement will enable the investigation of complex protein-membrane interactions in live cells or tissues.
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Affiliation(s)
- Bin Cai
- Department of Cancer Biology, University of Cincinnati, Cincinnati, OH 45267, USA; Research Center for Nano-Biomaterial, Analytical & Testing Center, Sichuan University, Chengdu 610064, China
| | - Luning Yu
- Department of Physics, University of Cincinnati, Cincinnati, OH 45221, USA
| | - Savanna R Sharum
- Department of Biochemistry, University of Illinois at Urbana-Champaign, Urbana, IL 61801, USA
| | - Kai Zhang
- Department of Biochemistry, University of Illinois at Urbana-Champaign, Urbana, IL 61801, USA.
| | - Jiajie Diao
- Department of Cancer Biology, University of Cincinnati, Cincinnati, OH 45267, USA.
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30
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Potassium channel selectivity filter dynamics revealed by single-molecule FRET. Nat Chem Biol 2019; 15:377-383. [PMID: 30833778 PMCID: PMC6430689 DOI: 10.1038/s41589-019-0240-7] [Citation(s) in RCA: 21] [Impact Index Per Article: 4.2] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 01/29/2018] [Accepted: 01/25/2019] [Indexed: 11/09/2022]
Abstract
Potassium (K) channels exhibit exquisite selectivity for conduction of K+ ions over other cations, particularly Na+. High-resolution structures reveal an archetypal selectivity filter (SF) conformation in which dehydrated K+ ions, but not Na+ ions, are perfectly coordinated. Using single-molecule FRET (smFRET), we show that the SF-forming loop (SF-loop) in KirBac1.1 transitions between constrained and dilated conformations as a function of ion concentration. The constrained conformation, essential for selective K+ permeability, is stabilized by K+ but not Na+ ions. Mutations that render channels nonselective result in dilated and dynamically unstable conformations, independent of the permeant ion. Further, while wild-type KirBac1.1 channels are K+ selective in physiological conditions, Na+ permeates in the absence of K+. Moreover, whereas K+ gradients preferentially support 86Rb+ fluxes, Na+ gradients preferentially support 22Na+ fluxes. This suggests differential ion selectivity in constrained versus dilated states, potentially providing a structural basis for this anomalous mole fraction effect.
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31
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Lin R, Wang Y. Automated smFRET microscope for the quantification of label-free DNA oligos. BIOMEDICAL OPTICS EXPRESS 2019; 10:682-693. [PMID: 30800508 PMCID: PMC6377893 DOI: 10.1364/boe.10.000682] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.6] [Reference Citation Analysis] [Abstract] [Track Full Text] [Subscribe] [Scholar Register] [Received: 10/03/2018] [Revised: 12/13/2018] [Accepted: 12/18/2018] [Indexed: 05/25/2023]
Abstract
Single-molecule FRET (smFRET) spectroscopy is a powerful tool for studying inhomogeneous dynamics in biological systems. However, because of the intrinsic variations that accompany the sample sizes, massive data sets are essential to extract statistically reliable information. In this aspect, a simple motorized stage and autofocusing modification can save time without the expense of a high-end automated microscope. In this report, we describe a simple and economical modification of a commercial inverted microscope with a manual stage to automate the data acquisition and measurement process. We collected 8000 images with a 100 ms exposure time in 1000 fields of view in approximately 13 min, where it would take more than 8 h by manual collection. We demonstrated the method with a DNA oligo quantification experiment. In this experiment, the measurement platform is a FRET signal from a dye-labeled DNA duplex containing unmatched base pairs. The target DNA replaces one of the strands because of the formation of a perfect duplex. This thermodynamically driven exchange reaction causes FRET to disappear, which correlated with the DNA concentration. The data are batch processed with the freeware ImageJ. These modifications are feasible and economical for general smFRET experiments.
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32
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Abstract
Cell division involves mechanical processes, such as chromosome transport and centrosome separation. Quantitative micromanipulation-based approaches have been central to dissecting the forces driving these processes. We highlight two biophysical assays that can be employed for such analyses. First, an in vitro "mini-spindle" assay is described that can be used to examine the collective mechanics of mitotic motor proteins cross-linking two microtubules. In the spindle, motor proteins (e.g., kinesin-5, kinesin-14, and dynein) can localize to overlapping microtubules that slide relative to each other, work as an ensemble, and equilibrate between cytoplasm and the microtubules. The "mini-spindle" assay can recapitulate these features and allows measurements of forces generated between adjacent microtubules and their dependence on filament orientation, sliding speed, overlap length, and motor protein density. Second, we describe a force-calibrated microneedle-based "whole-spindle" micromechanics assay. Microneedle-based micromanipulation can be a useful technique to examine cellular scale mechanics, but its use has been restricted by the difficulty in getting probes to penetrate the plasma membrane without disrupting cell physiology. As detailed here, the use of cell-free extracts prepared from metaphase-arrested Xenopus eggs can address this limitation. These micromanipulation studies also benefit from the use of frozen stocks of Xenopus egg extract. Together, these approaches can be used to decipher how micromechanics and biochemical activities ensure successful cell division.
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Affiliation(s)
- Yuta Shimamoto
- Center for Frontier Research, National Institute of Genetics, Mishima, Shizuoka, Japan.
| | - Tarun M Kapoor
- Laboratory of Chemistry and Cell Biology, The Rockefeller University, New York, NY, United States.
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33
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Börner R, Kowerko D, Hadzic MCAS, König SLB, Ritter M, Sigel RKO. Simulations of camera-based single-molecule fluorescence experiments. PLoS One 2018; 13:e0195277. [PMID: 29652886 PMCID: PMC5898730 DOI: 10.1371/journal.pone.0195277] [Citation(s) in RCA: 13] [Impact Index Per Article: 2.2] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 12/05/2017] [Accepted: 03/19/2018] [Indexed: 01/23/2023] Open
Abstract
Single-molecule microscopy has become a widely used technique in (bio)physics and (bio)chemistry. A popular implementation is single-molecule Förster Resonance Energy Transfer (smFRET), for which total internal reflection fluorescence microscopy is frequently combined with camera-based detection of surface-immobilized molecules. Camera-based smFRET experiments generate large and complex datasets and several methods for video processing and analysis have been reported. As these algorithms often address similar aspects in video analysis, there is a growing need for standardized comparison. Here, we present a Matlab-based software (MASH-FRET) that allows for the simulation of camera-based smFRET videos, yielding standardized data sets suitable for benchmarking video processing algorithms. The software permits to vary parameters that are relevant in cameras-based smFRET, such as video quality, and the properties of the system under study. Experimental noise is modeled taking into account photon statistics and camera noise. Finally, we survey how video test sets should be designed to evaluate currently available data analysis strategies in camera-based sm fluorescence experiments. We complement our study by pre-optimizing and evaluating spot detection algorithms using our simulated video test sets.
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Affiliation(s)
- Richard Börner
- Department of Chemistry, University of Zurich, Zurich, Switzerland
| | - Danny Kowerko
- Department of Computer Science, Chemnitz University of Technology, Chemnitz, Germany
| | | | - Sebastian L. B. König
- Department of Chemistry, University of Zurich, Zurich, Switzerland
- Department of Biochemistry, University of Zurich, Zurich, Switzerland
| | - Marc Ritter
- Department of Applied Computer and Biosciences, Mittweida University of Applied Sciences, Mittweida, Germany
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34
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Ma L, Li Y, Ma J, Hu S, Li M. Watching Three-Dimensional Movements of Single Membrane Proteins in Lipid Bilayers. Biochemistry 2018; 57:4735-4740. [PMID: 29619828 DOI: 10.1021/acs.biochem.8b00253] [Citation(s) in RCA: 1] [Impact Index Per Article: 0.2] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 11/30/2022]
Abstract
It is challenging to assess protein-membrane interactions because of the lack of appropriate tools to detect position changes of single proteins in the ∼4 nm range of biological membranes. We developed an assay recently, termed surface-induced fluorescence attenuation (SIFA). It is able to track both vertical and lateral dynamic motion of singly labeled membrane proteins in supported lipid bilayers. Similar to the FRET (fluorescence resonance energy transfer) principle, SIFA takes advantage of the energy transfer from a fluorophore to a light-absorbing surface to determine the distance at 2-8 nm away from the surface. By labeling a protein with a proper fluorophore and using graphene oxide as a two-dimensional quencher, we showed that SIFA is capable of monitoring three-dimensional movements of the fluorophore-labeled protein not only inside but also above the lipid bilayer atop the graphene oxide. Our data show that SIFA is a well-suited method to study the interplay between proteins and membranes.
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Affiliation(s)
- Li Ma
- Beijing National Laboratory for Condensed Matter Physics and CAS Key Laboratory of Soft Matter Physics, Institute of Physics , Chinese Academy of Sciences , Beijing 100190 , China.,School of Physical Sciences , University of Chinese Academy of Sciences , Beijing 100049 , China
| | - Ying Li
- Beijing National Laboratory for Condensed Matter Physics and CAS Key Laboratory of Soft Matter Physics, Institute of Physics , Chinese Academy of Sciences , Beijing 100190 , China.,School of Physical Sciences , University of Chinese Academy of Sciences , Beijing 100049 , China
| | - Jianbing Ma
- Beijing National Laboratory for Condensed Matter Physics and CAS Key Laboratory of Soft Matter Physics, Institute of Physics , Chinese Academy of Sciences , Beijing 100190 , China.,School of Physical Sciences , University of Chinese Academy of Sciences , Beijing 100049 , China
| | - Shuxin Hu
- Beijing National Laboratory for Condensed Matter Physics and CAS Key Laboratory of Soft Matter Physics, Institute of Physics , Chinese Academy of Sciences , Beijing 100190 , China.,School of Physical Sciences , University of Chinese Academy of Sciences , Beijing 100049 , China
| | - Ming Li
- Beijing National Laboratory for Condensed Matter Physics and CAS Key Laboratory of Soft Matter Physics, Institute of Physics , Chinese Academy of Sciences , Beijing 100190 , China.,School of Physical Sciences , University of Chinese Academy of Sciences , Beijing 100049 , China
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35
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Hayne CK, Yumerefendi H, Cao L, Gauer JW, Lafferty MJ, Kuhlman B, Erie DA, Neher SB. We FRET so You Don't Have To: New Models of the Lipoprotein Lipase Dimer. Biochemistry 2018; 57:241-254. [PMID: 29303250 PMCID: PMC5860654 DOI: 10.1021/acs.biochem.7b01009] [Citation(s) in RCA: 9] [Impact Index Per Article: 1.5] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/15/2023]
Abstract
Lipoprotein lipase (LPL) is a dimeric enzyme that is responsible for clearing triglyceride-rich lipoproteins from the blood. Although LPL plays a key role in cardiovascular health, an experimentally derived three-dimensional structure has not been determined. Such a structure would aid in understanding mutations in LPL that cause familial LPL deficiency in patients and help in the development of therapeutic strategies to target LPL. A major obstacle to structural studies of LPL is that LPL is an unstable protein that is difficult to produce in the quantities needed for nuclear magnetic resonance or crystallography. We present updated LPL structural models generated by combining disulfide mapping, computational modeling, and data derived from single-molecule Förster resonance energy transfer (smFRET). We pioneer the technique of smFRET for use with LPL by developing conditions for imaging active LPL and identifying positions in LPL for the attachment of fluorophores. Using this approach, we measure LPL-LPL intermolecular interactions to generate experimental constraints that inform new computational models of the LPL dimer structure. These models suggest that LPL may dimerize using an interface that is different from the dimerization interface suggested by crystal packing contacts seen in structures of pancreatic lipase.
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Affiliation(s)
- Cassandra K. Hayne
- Department of Biochemistry and Biophysics, University of North Carolina - Chapel Hill
| | - Hayretin Yumerefendi
- Department of Biochemistry and Biophysics, University of North Carolina - Chapel Hill
| | - Lin Cao
- Department of Biochemistry and Biophysics, University of North Carolina - Chapel Hill
| | - Jacob W. Gauer
- Department of Chemistry, University of North Carolina at Chapel Hill
| | - Michael J. Lafferty
- Department of Biochemistry and Biophysics, University of North Carolina - Chapel Hill
| | - Brian Kuhlman
- Department of Biochemistry and Biophysics, University of North Carolina - Chapel Hill
| | - Dorothy A. Erie
- Department of Chemistry, University of North Carolina at Chapel Hill
- Lineberger Comprehensive Cancer Center, University of North Carolina at Chapel Hill
| | - Saskia B. Neher
- Department of Biochemistry and Biophysics, University of North Carolina - Chapel Hill
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36
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Abstract
Human RAD51 promotes accurate DNA repair by homologous recombination and is involved in protection and repair of damaged DNA replication forks. The active species of RAD51 and related recombinases in all organisms is a nucleoprotein filament assembled on single-stranded DNA (ssDNA). The formation of a nucleoprotein filament competent for the recombination reaction, or for DNA replication support, is a delicate and strictly regulated process, which occurs through filament nucleation followed by filament extension. The rates of these two phases of filament formation define the capacity of RAD51 to compete with the ssDNA-binding protein RPA, as well as the lengths of the resulting filament segments. Single-molecule approaches can provide a wealth of quantitative information on the kinetics of RAD51 nucleoprotein filament assembly, internal dynamics, and disassembly. In this chapter, we describe how to set up a single-molecule total internal reflection fluorescence microscopy experiment to monitor the initial steps of RAD51 nucleoprotein filament formation in real-time and at single-monomer resolution. This approach is based on the unique, stretched-ssDNA conformation within the recombinase nucleoprotein filament and follows the efficiency of Förster resonance energy transfer (EFRET) between two DNA-conjugated fluorophores. We will discuss the practical aspects of the experimental setup, extraction of the FRET trajectories, and how to analyze and interpret the data to obtain information on RAD51 nucleation kinetics, the mechanism of nucleation, and the oligomeric species involved in filament formation.
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Glembockyte V, Cosa G. Redox-Based Photostabilizing Agents in Fluorescence Imaging: The Hidden Role of Intersystem Crossing in Geminate Radical Ion Pairs. J Am Chem Soc 2017; 139:13227-13233. [DOI: 10.1021/jacs.7b08134] [Citation(s) in RCA: 22] [Impact Index Per Article: 3.1] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/20/2022]
Affiliation(s)
- Viktorija Glembockyte
- Department of Chemistry and
Center for Self-Assembled Chemical Structures, McGill University, 801
Sherbrooke Street West, Montréal, Québec H3A 0B8, Canada
| | - Gonzalo Cosa
- Department of Chemistry and
Center for Self-Assembled Chemical Structures, McGill University, 801
Sherbrooke Street West, Montréal, Québec H3A 0B8, Canada
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Optimal Background Estimators in Single-Molecule FRET Microscopy. Biophys J 2017; 111:1278-1286. [PMID: 27653486 DOI: 10.1016/j.bpj.2016.07.047] [Citation(s) in RCA: 14] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 10/14/2015] [Revised: 06/21/2016] [Accepted: 07/26/2016] [Indexed: 11/23/2022] Open
Abstract
Single-molecule total internal reflection fluorescence (TIRF) microscopy constitutes an umbrella of powerful tools that facilitate direct observation of the biophysical properties, population heterogeneities, and interactions of single biomolecules without the need for ensemble synchronization. Due to the low signal/noise ratio in single-molecule TIRF microscopy experiments, it is important to determine the local background intensity, especially when the fluorescence intensity of the molecule is used quantitatively. Here we compare and evaluate the performance of different aperture-based background estimators used particularly in single-molecule Förster resonance energy transfer. We introduce the general concept of multiaperture signatures and use this technique to demonstrate how the choice of background can affect the measured fluorescence signal considerably. A new, to our knowledge, and simple background estimator is proposed, called the local statistical percentile (LSP). We show that the LSP background estimator performs as well as current background estimators at low molecular densities and significantly better in regions of high molecular densities. The LSP background estimator is thus suited for single-particle TIRF microscopy of dense biological samples in which the intensity itself is an observable of the technique.
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Maleki P, Budhathoki JB, Roy WA, Balci H. A practical guide to studying G-quadruplex structures using single-molecule FRET. Mol Genet Genomics 2017; 292:483-498. [PMID: 28150040 DOI: 10.1007/s00438-017-1288-2] [Citation(s) in RCA: 24] [Impact Index Per Article: 3.4] [Reference Citation Analysis] [Abstract] [Key Words] [MESH Headings] [Grants] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Received: 11/15/2016] [Accepted: 01/05/2017] [Indexed: 11/26/2022]
Abstract
In this article, we summarize the knowledge and best practices learned from bulk and single-molecule measurements to address some of the frequently experienced difficulties in single-molecule Förster resonance energy transfer (smFRET) measurements on G-quadruplex (GQ) structures. The number of studies that use smFRET to investigate the structure, function, dynamics, and interactions of GQ structures has grown significantly in the last few years, with new applications already in sight. However, a number of challenges need to be overcome before reliable and reproducible smFRET data can be obtained in measurements that include GQ. The annealing and storage conditions, the location of fluorophores on the DNA construct, and the ionic conditions of the experiment are some of the factors that are of critical importance for the outcome of measurements, and many of these manifest themselves in unique ways in smFRET assays. By reviewing these aspects and providing a summary of best practices, we aim to provide a practical guide that will help in successfully designing and performing smFRET studies on GQ structures.
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Affiliation(s)
- Parastoo Maleki
- Department of Physics, Kent State University, Kent, OH, 44242, USA
| | | | - William A Roy
- Department of Physics, Kent State University, Kent, OH, 44242, USA
| | - Hamza Balci
- Department of Physics, Kent State University, Kent, OH, 44242, USA.
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Maleki P, Ma Y, Iida K, Nagasawa K, Balci H. A single molecule study of a fluorescently labeled telomestatin derivative and G-quadruplex interactions. Nucleic Acids Res 2016; 45:288-295. [PMID: 27899628 PMCID: PMC5224478 DOI: 10.1093/nar/gkw1090] [Citation(s) in RCA: 11] [Impact Index Per Article: 1.4] [Reference Citation Analysis] [Abstract] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/02/2016] [Revised: 10/10/2016] [Accepted: 10/25/2016] [Indexed: 02/03/2023] Open
Abstract
The potential use of G-quadruplex (GQ) stabilizing small molecules as anti-cancer drugs has created a flurry of activity on various aspects of these molecules. Telomestatin and oxazole telomestatin derivatives (OTD) are some of the most prominent of such molecules, yet the underlying dynamics of their interactions with GQ and the extent of heterogeneities in these interactions are not known. We performed single molecule measurements to study binding kinetics, rotational freedom, and dwell time distributions of a Cy5-labeled OTD (L1Cy5–7OTD) as it interacted with several different GQ structures. Our measurements show that L1Cy5–7OTD dwells on more stable GQ for longer times and binds to such GQ with higher frequency. The dwell times showed a broad distribution, but were longer than a minute for a significant fraction of molecules (characteristic dwell time τ = 192 ± 15 s and τ = 98 ± 15 s for the more and less stable GQ, respectively). In addition, L1Cy5–7OTD might be able to bind to GQ in at least two different primary orientations and occasionally transition between these orientations. The dwell time in one of these orientations was significantly longer than that in the other one, suggesting different stabilities for different binding orientations.
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Affiliation(s)
- Parastoo Maleki
- Department of Physics, Kent State University, Kent, OH 44240, USA
| | - Yue Ma
- Department of Biotechnology and Life Science, Graduate School of Technology, Tokyo University of Agriculture and Technology, Koganei, Tokyo 184-8588, Japan
| | - Keisuke Iida
- Department of Biotechnology and Life Science, Graduate School of Technology, Tokyo University of Agriculture and Technology, Koganei, Tokyo 184-8588, Japan
| | - Kazuo Nagasawa
- Department of Biotechnology and Life Science, Graduate School of Technology, Tokyo University of Agriculture and Technology, Koganei, Tokyo 184-8588, Japan
| | - Hamza Balci
- Department of Physics, Kent State University, Kent, OH 44240, USA
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Tyrosine phosphorylation stimulates activity of human RAD51 recombinase through altered nucleoprotein filament dynamics. Proc Natl Acad Sci U S A 2016; 113:E6045-E6054. [PMID: 27671650 DOI: 10.1073/pnas.1604807113] [Citation(s) in RCA: 38] [Impact Index Per Article: 4.8] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/03/2023] Open
Abstract
The DNA strand exchange protein RAD51 facilitates the central step in homologous recombination, a process fundamentally important for accurate repair of damaged chromosomes, restart of collapsed replication forks, and telomere maintenance. The active form of RAD51 is a nucleoprotein filament that assembles on single-stranded DNA (ssDNA) at the sites of DNA damage. The c-Abl tyrosine kinase and its oncogenic counterpart BCR-ABL fusion kinase phosphorylate human RAD51 on tyrosine residues 54 and 315. We combined biochemical reconstitutions of the DNA strand exchange reactions with total internal reflection fluorescence microscopy to determine how the two phosphorylation events affect the biochemical activities of human RAD51 and properties of the RAD51 nucleoprotein filament. By mimicking RAD51 tyrosine phosphorylation with a nonnatural amino acid, p-carboxymethyl-l-phenylalanine (pCMF), we demonstrated that Y54 phosphorylation enhances the RAD51 recombinase activity by at least two different mechanisms, modifies the RAD51 nucleoprotein filament formation, and allows RAD51 to compete efficiently with ssDNA binding protein RPA. In contrast, Y315 phosphorylation has little effect on the RAD51 activities. Based on our work and previous cellular studies, we propose a mechanism underlying RAD51 activation by c-Abl/BCR-ABL kinases.
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42
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Single-molecule sorting of DNA helicases. Methods 2016; 108:14-23. [PMID: 27223403 DOI: 10.1016/j.ymeth.2016.05.009] [Citation(s) in RCA: 16] [Impact Index Per Article: 2.0] [Reference Citation Analysis] [Abstract] [Key Words] [Journal Information] [Subscribe] [Scholar Register] [Received: 03/25/2016] [Revised: 05/13/2016] [Accepted: 05/16/2016] [Indexed: 11/22/2022] Open
Abstract
DNA helicases participate in virtually all aspects of cellular DNA metabolism by using ATP-fueled directional translocation along the DNA molecule to unwind DNA duplexes, dismantle nucleoprotein complexes, and remove non-canonical DNA structures. Post-translational modifications and helicase interacting partners are often viewed as determining factors in controlling the switch between bona fide helicase activity and other functions of the enzyme that do not involve duplex separation. The bottleneck in developing a mechanistic understanding of human helicases and their control by post-translational modifications is obtaining sufficient quantities of the modified helicase for traditional structure-functional analyses and biochemical reconstitutions. This limitation can be overcome by single-molecule analysis, where several hundred surface-tethered molecules are sufficient to obtain a complete kinetic and thermodynamic description of the helicase-mediated substrate binding and rearrangement. Synthetic oligonucleotides site-specifically labeled with Cy3 and Cy5 fluorophores can be used to create a variety of DNA substrates that can be used to characterize DNA binding, as well as helicase translocation and duplex unwinding activities. This chapter describes "single-molecule sorting", a robust experimental approach to simultaneously quantify, and distinguish the activities of helicases carrying their native post-translational modifications. Using this technique, a DNA helicase of interest can be produced and biotinylated in human cells to enable surface-tethering for the single-molecule studies by total internal reflection fluorescence microscopy. The pool of helicases extracted from the cells is expected to contain a mixture of post-translationally modified and unmodified enzymes, and the contributions from either population can be monitored separately, but in the same experiment providing a direct route to evaluating the effect of a given modification.
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Goldfain AM, Garmann RF, Jin Y, Lahini Y, Manoharan VN. Dynamic Measurements of the Position, Orientation, and DNA Content of Individual Unlabeled Bacteriophages. J Phys Chem B 2016; 120:6130-8. [DOI: 10.1021/acs.jpcb.6b02153] [Citation(s) in RCA: 19] [Impact Index Per Article: 2.4] [Reference Citation Analysis] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 12/13/2022]
Affiliation(s)
| | | | - Yan Jin
- Department
of Chemistry and Biochemistry, University of California, Los Angeles, California 90095, United States
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Quantifying the Assembly of Multicomponent Molecular Machines by Single-Molecule Total Internal Reflection Fluorescence Microscopy. Methods Enzymol 2016; 581:105-145. [PMID: 27793278 PMCID: PMC5403009 DOI: 10.1016/bs.mie.2016.08.019] [Citation(s) in RCA: 15] [Impact Index Per Article: 1.9] [Reference Citation Analysis] [Abstract] [Key Words] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 01/24/2023]
Abstract
Large, dynamic macromolecular complexes play essential roles in many cellular processes. Knowing how the components of these complexes associate with one another and undergo structural rearrangements is critical to understanding how they function. Single-molecule total internal reflection fluorescence (TIRF) microscopy is a powerful approach for addressing these fundamental issues. In this article, we first discuss single-molecule TIRF microscopes and strategies to immobilize and fluorescently label macromolecules. We then review the use of single-molecule TIRF microscopy to study the formation of binary macromolecular complexes using one-color imaging and inhibitors. We conclude with a discussion of the use of TIRF microscopy to examine the formation of higher-order (i.e., ternary) complexes using multicolor setups. The focus throughout this article is on experimental design, controls, data acquisition, and data analysis. We hope that single-molecule TIRF microscopy, which has largely been the province of specialists, will soon become as common in the tool box of biophysicists and biochemists as structural approaches have become today.
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45
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Brown MS, Grubb J, Zhang A, Rust MJ, Bishop DK. Small Rad51 and Dmc1 Complexes Often Co-occupy Both Ends of a Meiotic DNA Double Strand Break. PLoS Genet 2015; 11:e1005653. [PMID: 26719980 PMCID: PMC4697796 DOI: 10.1371/journal.pgen.1005653] [Citation(s) in RCA: 62] [Impact Index Per Article: 6.9] [Reference Citation Analysis] [Abstract] [MESH Headings] [Grants] [Track Full Text] [Download PDF] [Figures] [Journal Information] [Subscribe] [Scholar Register] [Received: 08/26/2015] [Accepted: 10/15/2015] [Indexed: 11/19/2022] Open
Abstract
The Eukaryotic RecA-like proteins Rad51 and Dmc1 cooperate during meiosis to promote recombination between homologous chromosomes by repairing programmed DNA double strand breaks (DSBs). Previous studies showed that Rad51 and Dmc1 form partially overlapping co-foci. Here we show these Rad51-Dmc1 co-foci are often arranged in pairs separated by distances of up to 400 nm. Paired co-foci remain prevalent when DSBs are dramatically reduced or when strand exchange or synapsis is blocked. Super-resolution dSTORM microscopy reveals that individual foci observed by conventional light microscopy are often composed of two or more substructures. The data support a model in which the two tracts of ssDNA formed by a single DSB separate from one another by distances of up to 400 nm, with both tracts often bound by one or more short (about 100 nt) Rad51 filaments and also by one or more short Dmc1 filaments.
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Affiliation(s)
- M. Scott Brown
- Department of Molecular Genetics and Cell Biology, University of Chicago, Cummings Life Science Center, Chicago, Illinois, United States of America
| | - Jennifer Grubb
- Department of Radiation and Cellular Oncology, University of Chicago, Cummings Life Science Center, Chicago, Illinois, United States of America
| | - Annie Zhang
- Department of Radiation and Cellular Oncology, University of Chicago, Cummings Life Science Center, Chicago, Illinois, United States of America
| | - Michael J. Rust
- Department of Molecular Genetics and Cell Biology, University of Chicago, Cummings Life Science Center, Chicago, Illinois, United States of America
| | - Douglas K. Bishop
- Department of Molecular Genetics and Cell Biology, University of Chicago, Cummings Life Science Center, Chicago, Illinois, United States of America
- Department of Radiation and Cellular Oncology, University of Chicago, Cummings Life Science Center, Chicago, Illinois, United States of America
- * E-mail:
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47
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Ma L, Yang F, Zheng J. Application of fluorescence resonance energy transfer in protein studies. J Mol Struct 2014; 1077:87-100. [PMID: 25368432 DOI: 10.1016/j.molstruc.2013.12.071] [Citation(s) in RCA: 57] [Impact Index Per Article: 5.7] [Reference Citation Analysis] [Abstract] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 02/08/2023]
Abstract
Since the physical process of fluorescence resonance energy transfer (FRET) was elucidated more than six decades ago, this peculiar fluorescence phenomenon has turned into a powerful tool for biomedical research due to its compatibility in scale with biological molecules as well as rapid developments in novel fluorophores and optical detection techniques. A wide variety of FRET approaches have been devised, each with its own advantages and drawbacks. Especially in the last decade or so, we are witnessing a flourish of FRET applications in biological investigations, many of which exemplify clever experimental design and rigorous analysis. Here we review the current stage of FRET methods development with the main focus on its applications in protein studies in biological systems, by summarizing the basic components of FRET techniques, most established quantification methods, as well as potential pitfalls, illustrated by example applications.
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Affiliation(s)
- Linlin Ma
- Department of Physiology and Membrane Biology, University of California School of Medicine, Davis, CA 95616, USA ; Institute for Molecular Bioscience, The University of Queensland, St Lucia, QLD 4072, Australia
| | - Fan Yang
- Department of Physiology and Membrane Biology, University of California School of Medicine, Davis, CA 95616, USA
| | - Jie Zheng
- Department of Physiology and Membrane Biology, University of California School of Medicine, Davis, CA 95616, USA
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48
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Joo C, Ha T. Prism-type total internal reflection microscopy for single-molecule FRET. Cold Spring Harb Protoc 2012; 2012:2012/12/pdb.prot072041. [PMID: 23209134 DOI: 10.1101/pdb.prot072041] [Citation(s) in RCA: 3] [Impact Index Per Article: 0.3] [Reference Citation Analysis] [Abstract] [MESH Headings] [Track Full Text] [Journal Information] [Subscribe] [Scholar Register] [Indexed: 06/01/2023]
Abstract
Single-molecule (sm) fluorescence detection is a powerful method for studying biological events without time and population averaging. Förster (fluorescence) resonance energy transfer (FRET) is a spectroscopic technique in which the efficiency of energy transfer from donor to acceptor molecules is used to determine distances between molecules in the 30-80 Å range. Structural changes in biological molecules or relative motion between two interacting molecules can be detected by a change in FRET. A variant of smFRET is based on total internal reflection (TIR) microscopy, which can be set up in two ways, either using an oil-immersion (objective-type) or a water-immersion (prism-type) lens. This protocol describes the setup for prism-type TIR microscopy (excitation and emission), which has been a major imaging tool for smFRET since it was first introduced.
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Abstract
The last 15 years have witnessed the development of tools that allow the observation and manipulation of single molecules. The rapidly expanding application of these technologies for investigating biological systems of ever-increasing complexity is revolutionizing our ability to probe the mechanisms of biological reactions. Here, we compare the mechanistic information available from single-molecule experiments with the information typically obtained from ensemble studies and show how these two experimental approaches interface with each other. We next present a basic overview of the toolkit for observing and manipulating biology one molecule at a time. We close by presenting a case study demonstrating the impact that single-molecule approaches have had on our understanding of one of life's most fundamental biochemical reactions: the translation of a messenger RNA into its encoded protein by the ribosome.
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Affiliation(s)
- Ignacio Tinoco
- Department of Chemistry, University of California at Berkeley, Berkeley, California 94720, USA
| | - Ruben L. Gonzalez
- Department of Chemistry, Columbia University, New York, New York 10027, USA
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